7 Tips to Improve Your Western Blot
7 Tips to Improve Your Western Blot
Proteins are the molecular machinery of a cell, ensuring that every cell in the body can carry out its specific role. Scientists often wish to study proteins and one way to achieve this is by western blotting. This is a technique utilizing antibodies to identify a protein of interest from a mixture in a biological sample. Firstly, samples are separated by their molecular weight via gel electrophoresis. The proteins are then transferred to a solid membrane, either nitrocellulose or PVDF, followed by a blocking step to prevent antibodies non-specifically binding to the membrane. The protein of interest is probed for using a specific antibody and finally detected by a secondary antibody, often conjugated to a fluorophore. The membrane can then be scanned to visualize the protein of interest.
In the first few months of my PhD I was attempting a lot of western blots (something which I had only read about before, I mean, how hard could it be to get a nice single band on a clean blot?). It definitely wasn’t as easy as I expected – I was getting different results with every blot and my blots were ALWAYS full of speckles in the background.
This prompted me to do a lot of troubleshooting along the road to a perfect western, and here I will provide some tips on how to achieve this.
This cannot be emphasized heavily enough. Any dirt, dust or leftover gel on your plates or membrane WILL show up as background on your blots. And, because life is unfair, to me the background always seemed to coincide with where my bands should be therefore interfering with any subsequent quantification. One way to prevent background is to clean all plates and combs with either 1% SDS or 70% Ethanol followed by distilled water then wipe them dry. Moreover, make sure you are always wearing gloves and all other equipment e.g. forceps are clean, too.
Making your best gel
Sometimes, hours of work will have been put into preparing samples prior to carrying out the western blot; but if the gel is not properly prepared all your hard work might go to waste, which would be a massive shame!
Follow your recipes
Make sure your solutions are made up correctly and are at the right pH; the stacking gel is usually acidic at pH 6.8 and the separating gel is alkaline at pH 8.8.
The acrylamide percentage you use is proportional to how easy it is for proteins to move through the gel. If your protein of interest has a low molecular weight and your acrylamide percentage is too low, you risk it running off the gel. Before starting to prepare your gel, check the molecular weight of your proteins and decide which gel percentage is most appropriate.
Give the gels enough time to polymerize
Usually I prepare slightly more stacking gel and separating gel, therefore I can check the tube in which I have prepared the solutions to see if they have polymerized rather than manipulating the gel itself. I have found that making fresh 10% APS improved the polymerization of my gels, and it doesn’t take long to prepare. If a gel hasn’t properly polymerized this could lead to "wavy" bands. It is best to be patient at this stage!
Don’t let the gel dry out
After pouring the separating gel (ensuring you’ve left enough space for the stacking gel and comb) fill up the empty space with hydrated isopropanol whilst it is polymerizing. This will ensure that you achieve a flat line at the top of your separating gel and it won’t dry out! After the separating gel has polymerized, you can pour off your overlay solution, rinse well with ddH2O, before proceeding to pour your stacking gel on top. After you have prepared your gel, be sure to use it soon after and try to avoid letting it stand out at room temperature. If you have prepared gels in advance, there are ways to store these for future use. You can wrap them in paper soaked with SDS Running Buffer or pour some ddH2O on top of the gel and seal it in a ziplock bag, gels can be kept at 4°C for a couple of days. Should the gel start to shrink at the edges, it’s best just to start over!
I must repeat this to myself at least 100 times every time I’m carrying out a western blot! If the electrodes are plugged into the wrong positive/negative outlet the electric field will be reversed, and your samples would flow the wrong way out of the gel. This is a very easy mistake to make but takes hardly any time to double-check.
Transferring your proteins from gel to membrane
Proteins "Run to Red" or "Run to Anode"
A straightforward way to remember what goes where on your transfer sandwich is the phrase "run to red". If you think that proteins will be flowing from the gel into your membrane of choice, your gel needs to be on the negatively charged black side (cathode) and your membrane on the positively charged red side (anode), which is the same way that the current flows.
Handle your membrane properly!
Always wear gloves, make sure your forceps are clean and roll any bubbles out of your transfer sandwich. Like I said earlier, dust and dirt WILL show up on your blot! Air bubbles will prevent the protein migrating from gel to membrane in that area, to avoid this you can use a small roller or the edge of a plastic 50ml tube to push out any bubbles in your transfer sandwich.
Don’t let the transfer get too hot
Tank transfers can generate a lot of heat, which causes problems during transfer. To overcome this, ensure that the transfer buffer is pre-prepared, and it has had enough time to chill (it can always be prepared the day before). Often, blotters may come with ice packs which can be inserted into the tank, this will help to keep the temperature down. Make sure that your ice packs are filled and frozen prior to transferring your gels! Moreover, you can carry out the transfer on ice or in a cold room. Decreasing the voltage and decreasing the transfer time can also decrease the heat produced during transfer. You can try various times/voltages and figure out what works best for you and your protein!
Check if your transfer has worked
Before proceeding to blocking and immunodetection steps, it is useful to verify if your proteins have successfully transferred from the gel to the membrane. This can be quickly and easily achieved by a protein stain such as Ponceau-S. Through this method you will be also be able to tell if there were any problems during the transfer, such as an uneven transfer or air bubbles trapped in the transfer sandwich. You can take this time to mark the orientation of your membrane with pencil, especially useful if you don’t have a visible ladder!
Blocking and washing
Now, these might be the most important steps when it comes to minimizing the background on your membrane.
Blocking is a crucial step to increase the specificity of your antibodies by blocking unspecific antibody binding sites. To block my membranes, I use 5% BSA in TBS-T. A tip I have adopted is filtering this solution prior to using it, this removes any particulates that might show up as background. I normally block my membranes for 1 hour at room temperature. Other commonly used blocking agents include non-fat powdered milk.
It is essential to wash your membranes with TBS-T after each of your antibody incubation steps. I wash my membranes 3 times each for 5 minutes with gentle agitation. This will remove any unbound antibody, and therefore reduce background!
It is imperative to test a series of different concentrations of your antibodies and work out what gives you the best/cleanest band on your blot. Antibody manufacturers often provide this information on their data sheets. Too much antibody, and you may see excessive background on your blot, too little antibody and you might not see anything at all.
Make sure you include controls when testing your antibodies. For example, does the secondary antibody give rise to bands without any primary antibody present? If so, it might lead to a false positive result.
Be sure to double check the species reactivity of your antibodies and if they are compatible with your samples.
I use IRDye conjugated antibodies from LI-COR, which can be detected by their Odyssey® Imaging Systems. These dyes can be detected in either the 700nm or 800nm channels.
Troubleshoot your antibody concentrations – if you have too much secondary you risk unspecific binding to your membrane, and guess what, you’ll have a lot of background! Moreover, too much secondary might bind to your band of interest and saturate the signal, which you won’t then be able to quantify. I usually manually scan my membranes at every intensity on the Odyssey® scanner, therefore I can tell when the signal maxes out.
When detecting proteins in both the 700nm and 800nm channel, make sure that your secondaries are specifically detecting the correct protein in the correct channel. Highly cross adsorbed secondaries are recommended for detecting proteins in both channels, as they are less likely to cross react with each other.